Objective 1: Potentiate Notch-mediated ex vivo expansion of human HSPCs with hypoxia. To address whether Delta1 Notch ligand and hypoxia synergize to affect the growth and differentiation of human HSPCs, G-CSF mobilized human CD34+ cells from 4 healthy subjects were cultured for 21 days in the presence of cytokines in normoxia (21% O2) or hypoxia (1.5-2% O2) in vessels coated with fibronectin alone or combined with increasing concentrations of Delta1 (2.5, 5, 10 and 20 g/mL). Cultures were evaluated by flow cytometry, CFU assay and the repopulating potential of ex-vivo cultured cells was assessed by limiting dilution transplantation in sublethally irradiated NOD-Scid-IL2rynull (NSG) mice. A maximum 7.1-fold expansion of CD34+ cells and CFU was observed when cells were cultured with Delta1 under normoxic conditions. In contrast, CD34+ cells and CFU increased 18.3-fold with Delta1 in hypoxia. Notably, in hypoxic cultures containing Delta1, phenotypically defined HSCs (CD34+CD45A-CD90+CD49f+) increased 6.6-fold compared to uncultured cells, but these cells were undetectable under normoxic conditions. In limiting dilution transplantation studies, long-term repopulating HSCs (LT-HSCs) in uncultured CD34+ cells were measured at the expected frequency (1 in 7,706; 95% CI of 3,446 to 17,232). When analyzed at 3 months post-transplantation, a limited (1.5-fold) increase in LT-HSC frequency (1 in 5,090; 95% CI 2.456 to 10,550) was obtained in Delta1 normoxic cultures compared to uncultured cells. However, the frequency of LT-HSCs (1 in 1,586; 95% CI 680 to 3,701) was substantially increased in Delta1-hypoxic cultures (4.9-fold higher than uncultured cells and 4.2-fold higher than normoxic Delta1 cultures). This suggests that hypoxia synergistically and significantly potentiates Delta1-induced expansion of in-vivo repopulating potential of human HSPCs as compared to Delta1 induction alone. To test the applicability of this ex-vivo culture system for genetically manipulated human HSPCs, we subjected lentivirally labeled human HSPCs to a 21-day of Delta1-hypoxic culture system and subsequently evaluated their long-term engraftment in NSG mouse transplantation assays. 100,000 human mPB CD34+ cells were transduced with GFP- or NGFR-expressing LV vectors at a multiplicity of infection of 1 by using our standard transduction protocols. Following transduction, these cells were further ex-vivo cultured under hypoxic conditions at an optimized concentration of Delta1 (5 mcg/mL) for 21 days, and subsequently transplanted in sublethally irradiated NSG mice. 4 months after transplantation we found a significant increase in the percentage of both total and genetically marked CD45+ cells from the NSG mice transplanted with cells subjected to a 21-day of Delta1-hypoxic culture following LV transduction as compared to LV transduction alone. Notably, in NGFR-expression LV vector group, the percentage of both total and genetically marked CD45+ cells (CD45+NGFR+ cells) increased by 4-fold (p < 0.05, by one sided t-test). These data suggest that this ex-vivo culture system can be extended to lentivirally manipulated human HSPCs, and this finding provides a rationale to test this culture platform in human HSPCs subjected to CRISPR/Cas9-mediated genome editing. To further improve this expansion approach for clinical translation, studies are underway to characterize the mechanisms by which Notch and hypoxia signaling pathways synergize in human HSPCs. Objective 2: Transiently modulate the expression of key regulators of HSPC self-renewal using CRISPRa and CRISPRi. Ectopic expression of homeobox B4 (HOXB4), a transcription factor containing a highly conserved DNA-binding motif known as the homeodomain, has been found to enhance HSPC self-renewal in vitro and in vivo and has been suggested as an approach to expand HSPCs. However, overexpression of HOXB4 using retroviral vectors resulted in leukemia in large animal models and is considered too risky to for any clinical applications. Transient induction of HOXB4 expression in HSPCs could represent an alternative approach to harnessing the power of HOXB4 on HSPCs. For instance, a soluble recombinant HOXB4 protein induced rapid ex vivo expansion of transduced HSPCs, thereby avoiding the use of integrating retroviral vectors while benefiting from the self-renewal capacity of HOXB4. However, the short half-life of the recombinant protein has proven a practical hurdle and alternative approaches are thus needed. In recent studies, loss-of-function somatic mutations in key epigenetic regulators, including DNMT3A, TET2 and ASXL1, have also been shown to confer a proliferative advantage on HSPCs, resulting in age-related clonal hematopoiesis (CH). Analysis of genetic mutations in these genes in mouse models have suggested their association with enhanced HSPC self-renewal. Similar to HOXB4, permanent inactivation of DNMT3A, TET2 or ASXL1 may also predispose to the development of malignancies in cooperation with secondary mutations that drive disease phenotype. Therefore, we hypothesized that transient inactivation of DNMT3A, TET2 or ASXL1 activity might allow HSPC expansion in vitro without increasing susceptibility to malignancies. To investigate this possibility, CRISPR/Cas9-based transcriptional activation (CRISPRa) and inhibition (CRISPRi) are developed to transiently induce or repress, respectively, the endogenous expression of select target genes. CRISPRa and CRISPRi utilize synthetic single guide RNAs (sgRNAs) to direct to a gene-of-interest a nuclease-inactive dead Cas9 (dCas9) fused to a transcriptional activator (e.g. VP64, P65) or repressor (e.g. KRAB) domain. Current approaches to CRISPRa and CRISPRi rely on electroporation of bulky exogenous plasmid DNA or on transduction of HSPCs with lentiviral vectors to deliver the required Cas9/sgRNA activator and repressor components. However, use of plasmids results in pronounced cytotoxicity to HSPCs and lentiviral transduction introduces the risk of insertional mutagenesis and, due to vector integration, is not readily amenable to the induction of transient changes to the cellular developmental program required for successful HSPC expansion. In contrast, prior CRISPR work using a nuclease-active Cas9 for genome editing of HSPCs demonstrated that the Cas9/sgRNA system delivered by electroporation as a ribonucleoprotein (RNP) complex is more effective than plasmid-based reagents and results in only minimal toxicity in HSPCs. Importantly, RNPs are also subject to the intrinsic proteasomal activity of the cell and thus display a limited intracellular half-life, a feature of interest for transient gene activation or repression. In FY19, we have primarily developed approaches to recombinantly produce and purify dCas9-activator or dCas9-repressor domain fusion proteins. However, yields are typically low as most proteins exceed 160 kDa. To overcome this limitation, we have used of a modified approach whereby the activator and repressor domains are separated from dCas9. This system is based on the incorporation of MS2 RNA aptamers at two loops of the sgRNA that are exposed in the ternary Cas9 complex. These aptamers can recruit MS2-activator (MS2-P65-HCF1, known as MPH) or MS2-repressor (MS2-KRAB) domains into the Cas9-sgRNA complex at the target gene locus. Unlike large Cas9 fusion proteins, the MPH and MS2-KRAB domains are much smaller (56 kDa and 26 kDa, respectively), and can therefore be produced and purified more easily. Work is underway to transiently activate or repress gene expression in HSPCs with this system, as a novel approach to induce ex vivo HSPC expansion.